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The circadian regulation of eclosion in
Drosophila melanogaster
Die zeitliche Steuerung des Adultschlupfes in
Drosophila melanogaster
Doctoral thesis for a doctoral degree
at the Graduate School of Life Sciences,
Julius-Maximilians-Universität Würzburg,
Section Integrative Biology
Submitted by
Franziska Ruf
from Heilbronn
Würzburg, 2016
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The present work was accomplished at the Department of
Neurobiology and Genetics at the
University of Würzburg.
Submitted on: 24.06.2016
Members of the Promotionskomitee:
Chairperson: Prof. Dr. Jörg Schultz
Primary Supervisor: Prof. Dr. Christian Wegener
Supervisor (Second): Prof. Dr. Wolfgang Rössler
Supervisor (Third): Prof. Dr. Sakiko Shiga
Date of Public Defence: …………………………………………….…………
Date of Receipt of Certificates: …………………………………………
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Table of contents
Table of contents 1
Summary 7
Zusammenfassung 9
Introduction 13
1. The clock – an endogenous timekeeping system 13
2. The circadian clock of Drosophila melanogaster 14
2.1 The molecular clock 15
2.2 The clock network 17
2.3 Entrainment pathways of the circadian clock 18
2.4 The clock-related roles of pigment-dispersing factor (PDF)
20
3. Eclosion in Drosophila melanogaster 20
3.1 Hormonal regulation of eclosion 23
3.2 Circadian regulation of eclosion 25
3.3 Peptidergic regulation of eclosion in Drosophila
melanogaster 25
4. Aim of the present dissertation research 27
Materials and Methods 29
1. Eclosion assays 29
1.1 Würzburg Eclosion Monitor (WEclMon) 29
1.2 Eclosion assays under light entrainment 30
1.3 Eclosion assays under temperature entrainment 30
1.4 Eclosion assays after optogenetic activation via
channelrhodopsin 31
1.5 Eclosion assays under natural condition 31
1.6 Analysis of rhythmicity 32
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2. Molecular analysis 32
2.1 Analysis of torso expression via Reverse Transcription
Polymerase-Chain-
Reaction (RT-PCR) 32
2.2 Analysis of the temporal expression of Ptth mRNA via
quantitative Real-time
Polymerase-Chain-Reaction (qPCR) 34
3. Fly strains 38
Chapter I. Development of a new eclosion monitoring system
(WEclMon) 41
1. Introduction 41
2. Results 43
2.1 WEclMon 43
2.2 FiJi macro 45
2.3 Comparison of eclosion profiles recoreded in the WEclMon
system and the
TriKinetics system 46
2.4 The wildtypes Canton S and Lindelbach under temperature
entrainment 46
3. Discussion 53
Chapter II. Eclosion rhythms under natural conditions 54
1. Introduction 54
2. Results 57
2.1 Humidity entrainment 60
2.2 Statistical modelling 61
2.3 Eclosion profiles of Canton S under natural conditions
62
2.4 Eclosion profiles of the per01 mutant under natural
conditions 68
2.5 Eclosion profiles of the pdf01 mutant under natural
conditions 73
2.6 Eclosion profiles of the han5304 mutant under natural
conditions 78
2.7 Comparison of the rhythmicity indices between the genotypes
under different
conditions 83
2.8 Statistical modelling of the factors affecting the daily
eclosion pattern 85
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2.9 Theoretical statistical modelling 86
2.10 Eclosion rhythms of the wildtype Hubland under laboratory
LD conditions 94
3. Discussion 95
Chapter III. The role of PTTH in the circadian timing of
eclosion and eclosion
rhythmicity 101
1. Introduction 101
2. Results 104
2.1 Eclosion rhythms under light entrainment after ablation of
PTTH neurons 104
2.2 Eclosion rhythms under temperature entrainment after
ablation of PTTH
neurons 104
2.3 Eclosion rhythms under temperature entrainment after
silencing of PTTH
neurons 104
2.4 Eclosion rhythms under light entrainment after knockdown of
the PDF
receptor in PTTH neurons 111
2.5 Eclosion rhythms under light entrainment after knockdown of
the sNPF
receptor in PTTH neurons 111
2.6 Eclosion rhythms under light entrainment after knockdown of
torso in the
prothoracic gland 116
2.7 Eclosion rhythms under light entrainment after knockdown of
torso in the
brain 116
2.8 Analysis of torso expression pattern 121
2.9 Analysis of the temporal expression of Ptth mRNA 122
3. Discussion 123
Chapter IV. The role of CCAP in the circadian timing of eclosion
and eclosion
rhythmicity 126
1. Introduction 126
2. Results 128
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2.1 Eclosion rhythms under light entrainment after ablation of
CCAP neurons 128
2.2 Eclosion rhythms under light entrainment after silencing of
CCAP neurons 128
2.3 Eclosion rhythms of the CCAPexc7 mutant under light
entrainment 128
2.4 Eclosion rhythms of the CCAPexc7 mutant under temperature
entrainment 128
2.5 Eclosion rhythms of the CCAPexc7 mutant under natural
conditions 137
3. Discussion 145
Chapter V. Screen for candidate peptides regulating eclosion
behavior and
rhythmicity 147
1. Introduction 147
2. Results 149
2.1 Ecdysis-triggering hormone (ETH) 149
2.2 Apterous (Ap) 153
2.3 Capability (Capa) 153
2.4 Corazonin (Crz) 157
2.5 Diuretic hormone 31 (DH31) 163
2.6 Diuretic hormone 44 (DH44) 163
2.7 Dromyosuppressin (DMS) 163
2.8 Eclosion hormone (EH) 172
2.9 Hugin (hug) 175
2.10 Mai316 175
2.11 Myoinhibitory Peptide (MIP) 178
2.12 Neuropeptide F (NPF) 178
2.13 Pigment dispersing factor (PDF) 182
2.14 Phantom (Phm) / Ecdysone 182
2.15 Prothoracicotropic hormone (PTTH) 182
2.16 Short Neuropeptide F (sNPF) 186
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2.17 Adipokinetic hormone (AKH) 186
2.18 Allatostatin A (AstA) 186
2.19 Tyrosine decarboxylase (Tdc) / Octopamine 191
2.20 Tyrosine hydroxylase (TH) / Dopamine 192
2.21 Tryptophan hydroxylase (TRH) / Serotonin 192
2.22 Summary 197
3. Discussion 199
Synopsis 202
Appendix 208
References 233
Publications 251
Curriculum vitae Fehler! Textmarke nicht definiert.
Acknowledgments 255
Affidavit 257
Eidesstattliche Erklärung 257
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Summary
7
Summary
Eclosion is the emergence of an adult insect from the pupal case
at the end of development.
In the fruit fly Drosophila melanogaster, eclosion is a
circadian clock-gated event and is
regulated by various peptides. When studied on the population
level, eclosion reveals a clear
rhythmicity with a peak at the beginning of the light-phase that
persists also under constant
conditions. It is a long standing hypothesis that eclosion
gating to the morning hours with
more humid conditions is an adaption to reduce water loss and
increase the survival. Eclosion
behavior, including the motor pattern required for the fly to
hatch out of the puparium, is
orchestrated by a well-characterized cascade of peptides. The
main components are ecdysis-
triggering hormone (ETH), eclosion hormone (EH) and crustacean
cardioactive peptide (CCAP).
The molt is initiated by a peak level and pupal ecdysis by a
subsequent decline of the
ecdysteroid ecdysone. Ecdysteroids are produced by the
prothoracic gland (PG), an endocrine
tissue that contains a peripheral clock and degenerates shortly
after eclosion. Production and
release of ecdysteroids are regulated by the prothoracicotropic
hormone (PTTH).
Although many aspects of the circadian clock and the peptidergic
control of the eclosion
behavior are known, it still remains unclear how both systems
are interconnected. The aim of
this dissertation research was to dissect this connection and
evaluate the importance of
different Zeitgebers on eclosion rhythmicity under natural
conditions.
Potential interactions between the central clock and the
peptides regulating ecdysis motor
behavior were evaluated by analyzing the influence of CCAP on
eclosion rhythmicity. Ablation
and silencing of CCAP neurons, as well as CCAP null-mutation did
not affect eclosion
rhythmicity under either light or temperature entrainment nor
under natural conditions.
To dissect the connection between the central and the peripheral
clock, PTTH neurons were
ablated. Monitoring eclosion under light and temperature
entrainment revealed that eclosion
became arrhythmic under constant conditions. However, qPCR
expression analysis revealed
no evidence for cycling of Ptth mRNA in pharate flies. To test
for a connection with pigment-
dispersing factor (PDF)-expressing neurons, the PDF receptor
(PDFR) and short neuropeptide
F receptor (sNPFR) were knocked down in the PTTH neurons.
Knockdown of sNPFR, but not
PDFR, resulted in arrhythmic eclosion under constant darkness
conditions. PCR analysis of the
PTTH receptor, Torso, revealed its expression in the PG and the
gonads, but not in the brain
or eyes, of pharate flies. Knockdown of torso in the PG lead to
arrhythmicity under constant
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Summary
8
conditions, which provides strong evidence for the specific
effect of PTTH on the PG. These
results suggest connections from the PDF positive lateral
neurons to the PTTH neurons via
sNPF signaling, and to the PG via PTTH and Torso. This
interaction presumably couples the
period of the peripheral clock in the PG to that of the central
clock in the brain.
To identify a starting signal for eclosion and possible further
candidates in the regulation of
eclosion behavior, chemically defined peptidergic and aminergic
neurons were
optogenetically activated in pharate pupae via ChR2-XXL. This
screen approach revealed two
candidates for the regulation of eclosion behavior:
Dromyosuppressin (DMS) and myo-
inhibitory peptides (MIP). However, ablation of DMS neurons did
not affect eclosion
rhythmicity or success and the exact function of MIP must be
evaluated in future studies.
To assess the importance of the clock and of possible Zeitgebers
in nature, eclosion of the
wildtype Canton S and the clock mutant per01 and the PDF
signaling mutants pdf01 and han5304
was monitored under natural conditions. For this purpose, the
Würzburg eclosion monitor
(WEclMon) was developed, which is a new open monitoring system
that allows direct
exposure of pupae to the environment. A general decline of
rhythmicity under natural
conditions compared to laboratory conditions was observed in all
tested strains. While the
wildtype and the pdf01 and han5304 mutants stayed weakly
rhythmic, the per01 mutant flies
eclosed mostly arrhythmic. PDF and its receptor (PDFR encoded by
han) are required for the
synchronization of the clock network and functional loss can
obviously be compensated by a
persisting synchronization to external Zeitgebers. The loss of
the central clock protein PER,
however, lead to a non-functional clock and revealed the
absolute importance of the clock for
eclosion rhythmicity. To quantitatively analyze the effect of
the clock and abiotic factors on
eclosion rhythmicity, a statistical model was developed in
cooperation with Oliver Mitesser
and Thomas Hovestadt. The modelling results confirmed the clock
as the most important
factor for eclosion rhythmicity. Moreover, temperature was found
to have the strongest effect
on the actual shape of the daily emergence pattern, while light
has only minor effects. Relative
humidity could be excluded as Zeitgeber for eclosion and
therefore was not further analyzed.
Taken together, the present dissertation identified the so far
unknown connection between
the central and peripheral clock regulating eclosion.
Furthermore, a new method for the
analysis of eclosion rhythms under natural conditions was
established and the necessity of a
functional clock for rhythmic eclosion even in the presence of
multiple Zeitgebers was shown.
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Zusammenfassung
9
Zusammenfassung
Der Schlupf adulter Fliegen aus dem Puparium wird in der
Taufliege Drosophila melanogaster
zum einen von der inneren Uhr und zum anderen von Peptiden
gesteuert. Beobachtet man
den Schlupf auf der Populationsebene, lässt sich erkennen, dass
die meisten Fliegen zu Beginn
der Lichtphase schlüpfen. Diese Rhythmizität im Schlupfverhalten
von Fliegenpopulationen
hält auch unter konstanten Bedingungen an. Seit langer Zeit wird
angenommen, dass der
Schlupf am Morgen eine Anpassung an feuchte Bedingungen ist,
wodurch der Wasserverlust
verringert und die Überlebenswahrscheinlichkeit erhöht werden
könnte. Das stereotype
motorische Schlupfverhalten, mit dem sich die Fliege aus der
Puppenhülle befreit, wird durch
das gut untersuchte Zusammenspiel zahlreicher Peptide gesteuert.
Die wichtigsten Peptide
sind hierbei das ecdysis-triggering hormone (ETH), das
Schlupfhormon (EH) und das
crustacean cardioactive peptide (CCAP). Wie bei jedem Schlupf
wird die Häutung durch eine
stark erhöhte Produktion des Ecdysteroids Ecdyson ausgelöst. Der
anschließende Abfall der
Ecdyson-Titer löst dann den Adultschlupf aus. Ecdysteroide
werden in der Prothorakaldrüse
(PD) gebildet, die eine periphere Uhr besitzt und kurz nach dem
Adultschlupf zurückgebildet
wird. Das prothorakotrope Hormon (PTTH) reguliert sowohl die
Produktion als auch die
Freisetzung der Ecdysteroide aus der PD.
Obwohl bereits viel über den Aufbau und die Funktionsweise der
inneren Uhr und der
Kontrolle des Adultschlupfes durch Peptide bekannt ist, weiß man
bisher nicht, wie beide
Systeme miteinander interagieren. Das Hauptziel der vorliegenden
Arbeit war es, einerseits
diese Verbindung zu untersuchen und andererseits die Gewichtung
verschiedener Zeitgeber
für den Adultschlupf unter natürlichen Bedingungen zu
bewerten.
Um eine mögliche Verbindung zwischen der zentralen Uhr und den
Peptiden, die das
motorische Verhalten während des Schlupfes steuern, zu
untersuchen, wurde der Einfluss von
CCAP auf die Schlupfrhythmik betrachtet. Hierzu wurden die
CCAP-exprimierenden Neurone
genetisch ablatiert oder elektrisch stillgelegt, sowie
zusätzlich eine CCAP-defiziente Mutante
getestet. Weder unter künstlichen Licht- oder Temperaturzyklen,
noch unter natürlichen
Bedingungen wurden Effekte auf den Schlupfrhythmus bei
veränderter CCAP Verfügbarkeit
beobachtet.
Die Verbindung zwischen der zentralen und der peripheren Uhr der
PD wurde untersucht,
indem die PTTH-exprimierenden Neurone in Fliegen ablatiert
wurden. Dies führte sowohl
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Zusammenfassung
10
unter konstanten Licht- als auch Temperaturbedingungen zu
arrhythmischem Schlupf der
Populationen. Die Analyse der Expression von Ptth mRNA mittels
qPCR lieferte keine Hinweise
auf eine zyklische Regulation des Ptth Transkripts in pharaten
Tieren. Um eine Verbindung zu
pigment-dispersing factor (PDF)-exprimierenden Uhrneuronen
nachzuweisen, wurden die
Rezeptoren von PDF (PDFR) und dem short Neuropeptide F (sNPFR)
in den PTTH- Neuronen
herunterreguliert. Nur der Verlust von sNPFR führte unter
konstanten Bedingungen zu
arrhythmischem Schlupf. RT-PCR-Analyse der mRNA Expression des
Rezeptors von PTTH,
Torso, ergab, dass torso mRNA in pharaten Fliegen nur in der PD
und in den Gonaden
exprimiert wird, nicht jedoch im Gehirn. Das Herrunterregulieren
der torso mRNA in der PD
führte unter konstanten Bedingungen zu arrhythmischem Schlupf
und lieferte deutliche
Hinweise zur spezifischen Funktion von PTTH in der PD. Diese
Ergebnisse zeigen eine sNPF-
vermittelte Verbindung zwischen den PDF-positiven lateralen
Neuronen und den PTTH-
Neuronen, welche über PTTH und Torso weiter bis in die PD
reicht. Durch diese Verbindung
wird vermutlich die Periode der peripheren Uhr in der PD an die
Periode der zentralen Uhr im
Gehirn angepasst.
Um ein Startsignal für den Adultschlupf und weitere mögliche
Kandidaten, die eine Rolle in
der Steuerung des Schlupfes spielen, zu identifizieren, wurden
chemisch definierte kleine
Gruppen peptiderger und aminerger Neurone optogenetisch durch
das Kanalrhodopsin ChR2-
XXL aktiviert. In dieser Testreihe wurden Dromyosuppressin (DMS)
und myoinhibitorisches
Peptid (MIP) als mögliche Kandidaten ermittelt. Eine Ablation
der DMS-Neurone hatte jedoch
keine Auswirkungen auf Schlupfrhythmik und -erfolg. Die genaue
Funktion von MIP sollte in
zukünftigen Experimenten untersucht werden.
Um die Gewichtung der Uhr und möglicher Zeitgeber für das
natürliche Verhalten zu
bestimmen, wurde der Schlupf des Wildtyps Canton S, der
Uhrmutante per01 sowie der PDF-
Signalwegsmutanten pdf01 und han5304 (han codiert für den PDFR)
unter natürlichen
Bedingungen beobachtet. Hierfür wurde ein neues und offenes
Aufzeichnungssystem
entwickelt: der Würzburger Schlupfmonitor (WEclMon), der einen
direkten Kontakt der
Puppen mit den sie umgebenden abiotischen Bedingungen
ermöglicht.
Im Vergleich zu Laborbedingungen war die Rhythmizität des
Schlupfes unter natürlichen
Bedingungen in allen getesteten Fliegenlinien weniger
ausgeprägt. Während der Wildtyp
sowie die pdf01 und han5304 Mutanten weiterhin schwach
rhythmisch schlüpften, schlüpfte die
per01 Mutante hauptsächlich arrhythmisch. Das Zusammenspiel
zwischen PDF und seinem
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Zusammenfassung
11
Rezeptor synchronisiert das Uhrnetzwerk, und der Verlust dieser
Interaktion kann durch
tägliches neues Ausrichten an den Zeitgebern ausgeglichen
werden. Der Verlust des
Uhrproteins PER unterbindet jedoch die komplette
Funktionsfähigkeit der Uhr. Dadurch wird
die Notwendigkeit der Uhr für einen rhythmischen Schlupf
unterstrichen. Um den Einfluss der
Uhr und abiotischer Faktoren auf den Schlupfrhythmus zu
untersuchen, wurde im Rahmen
einer Kooperation mit Oliver Mitesser und Thomas Hovestadt ein
statistisches Modell
entwickelt. Die Ergebnisse der Modellierung unterstützen die
Hypothese, dass die Uhr der
wichtigste Faktor für einen rhythmischen Schlupf auch unter
Zeitgeber-Bedingungen ist. Die
Umgebungstemperatur übt hingegen den stärksten Einfluss auf die
Form des täglichen
Schlupfmusters aus, während Licht hier nur einen schwachen
Einfluss hat. Es konnte gezeigt
werden, dass sich relative Luftfeuchtigkeit nicht als Zeitgeber
für den Schlupf eignet, weshalb
sie in weiteren Untersuchungen nicht berücksichtigt wurde.
Zusammenfassend lässt sich sagen, dass mit der vorliegenden
Arbeit die Verbindung zwischen
der zentralen und peripheren Uhr in der Steuerung des Schlupfes
identifiziert werden
konnten, die bisher nicht bekannt war. Außerdem wurde eine neue
Methode der
Untersuchung des Adultschlupfes unter natürlichen Bedingungen
etabliert und die
Notwendigkeit einer intakten Uhr für einen rhythmischen
Adultschlupf selbst in Anwesenheit
mehrerer Zeitgeber konnte herausgestellt werden.
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Introduction
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Introduction
The clock – an endogenous timekeeping system
The rotation of the earth around its own axis and around the sun
leads to periodic changes in
the environment that can occur daily, monthly or annually. To
adapt physiological and
biochemical processes as well as their behavior to these
changes, all known organisms, from
bacteria to mammals, evolved endogenous timekeeping systems, so
called endogenous
clocks. These clocks allow the anticipation of periodic changes
in the environment and are
therefore thought to increase the fitness of the individual.
Rhythms can be found at different
levels of organization, ranging from cells over tissues to
individuals and even populations
(Saunders, 2002; Zordan et al., 2009). Endogenous clocks that
oscillate with rhythms of
approximately 24 hours are called circadian (from the Latin
circa = around and dianus = daily)
and represent the majority of clocks.
Among the first reports on rhythmic behaviors are studies of the
French scientist De Mairan
on the daily leaf movements of Mimosa pudica from 1729. Around
200 years later, one of the
first experimental studies on rhythmic animal behavior was on
the eclosion of fly (Pegomyia
hyoscyami) and moth (Ephestia kühniella) populations in the
morning (Bremer, 1926) and an
underlying endogenous timekeeping system was later described for
the fruit fly Drosophila
melanogaster (Bünning, 1935; Kalmus, 1935; Kalmus, 1938; Kalmus,
1940). Early works on the
properties of biological clocks by Pittendrigh, Zimmerman, Bruce
and others in the 1950-
1960ies lead to the following defining characteristics (reviewed
in Saunders, 2002):
1) clocks are able to entrain to so called Zeitgebers, which
means they synchronize their
period to environmental variables
2) clocks display rhythms that persist also under constant
conditions in the absence of
cues from the environment; they are then free-running
3) the free-running period is close to the period of the
environment to which the rhythm
is entrained
4) clocks are temperature-compensated; while most biochemical
and physiological
processes increase their rate with increasing temperatures, the
period of the clock is
not affected by temperature changes.
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Introduction
14
While the most prominent Zeitgebers are daily changes in light
and temperature, also
humidity, social contacts, food availability, tidal changes and
other environmental influences
can entrain the clock. Besides the central clock, which is
located in the brain, many body
tissues have peripheral clocks that oscillate autonomously but
are coupled with the central
clock in animals (Figure 1).
Figure 1: Schematic model of the clock system The central clock
receives input from Zeitgebers like light, temperature or food
availability, and thereby entrains its period to the period of the
Zeitgeber. Peripheral clocks also receive input from Zeitgebers,
but may also be coupled to the central clock. Alone or together
with peripheral clocks, the central clock generates different
outputs, for example on the physiological or behavioral level.
The circadian clock of Drosophila melanogaster
Drosophila melanogaster is one of the most used model organisms
for research on circadian
clocks. Early chronobiologists like Bünning and Pittendrigh
studied Drosophila melanogaster
and D. pseudoobscura to characterize the general properties of
clocks (reviewed in Saunders,
2002). The first clock gene, period, was discovered in
Drosophila (Konopka and Benzer, 1971).
Many fly behaviors have been shown to be under circadian
regulation, for example locomotor
activity (Roberts, 1956), eclosion (Bremer, 1926; Pittendrigh,
1954), oviposition (Allemand,
1976a; Allemand, 1976b), sensitivity to olfactory (Krishnan et
al., 1999) and gustatory
stimulation (Chatterjee et al., 2010), courtship behavior
(Hardeland, 1972; Fujii et al., 2007)
and learning and memory (Lyons and Roman, 2009).
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Introduction
15
2.1 The molecular clock
The molecular clock of Drosophila melanogaster consists of two
interlocked transcriptional
feedback loops that comprise the basic generator of the
circadian clock (Figure 2). At the
center of these feedback loops are the clock proteins CLOCK
(CLK) and CYCLE (CYC) that form
heterodimers and act as transcriptional activators (Hogenesch et
al., 1998). They induce
expression of target genes by binding to target sequences, so
called E boxes, located in the
promotor regions (Allada et al., 1998; Darlington et al., 1998;
Rutila et al., 1998). In the first
feedback loop, CLK-CYC activates the transcription of the clock
genes period (per) and timeless
(tim). PER and TIM proteins accumulate in the cytoplasm and
enter the nucleus as
heterodimers (Curtin et al., 1995; Gekakis et al., 1995; Saez
and Young, 1996; Zeng et al., 1996;
Kloss et al., 1998; Price et al., 1998). In the nucleus, the
complex inhibits the transcription of
their own genes through an interaction of PER with CLK, which
disables the CLK-CYC
heterodimer from binding to the E boxes (Darlington et al.,
1998). The mRNA levels of per and
tim peak at the end of the light phase (Hardin et al., 1990;
Sehgal et al., 1995) and their
proteins accumulate in the nucleus only during the dark phase
(Hardin et al., 1990). This is due
to the action of the blue-light receptor protein Cryptochrome
(CRY) which is activated by light
(Emery et al., 1998; Stanewsky et al., 1998). Activated CRY
binds to TIM and leads to its
proteasome-mediated degradation (Ceriani et al., 1999; Naidoo et
al., 1999; Busza et al.,
2004). Without TIM, PER is phosphorylated by the kinase
Double-Time (DBT), which leads to
PER degradation (Kloss et al., 1998; Price et al., 1998; Kloss
et al., 2001). The process of light-
mediated degradation of TIM and PER synchronizes the circadian
clock with the environment.
The kinase Shaggy (SGG) stabilizes CRY through binding and saves
TIM from degradation by
an unknown mechanism (Martinek et al., 2001; Stoleru et al.,
2007). In the first feedback loop,
PER and TIM can accumulate in the nucleus and inhibit CLK-CYC
only during the night, while
during the day, CLK-CYC can promote per and tim transcription
and restart the transcriptional
loop.
In a second feedback loop, CLK-CYC induces the transcription of
the clock genes vrille (vri)
(Blau and Young, 1999) and Par domain protein 1 (Pdp1) (McDonald
and Rosbash, 2001; Ueda
et al., 2002) by binding to their E-boxes. The proteins then
bind to V/P boxes in the promotor
region of clk (Cyran et al., 2003; Glossop et al., 2003) and
regulate its transcription: while VRI
is acting as a repressor for clk expression (Glossop et al.,
2003), PDP1 is acting as an activator
(Cyran et al., 2003). With clockwork orange (cwo) another
CLK-CYC activated clock gene was
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Introduction
16
discovered, which acts as a transcriptional repressor of CLK
target genes by binding to their E
boxes (Figure 3) (Kadener et al., 2007; Lim et al., 2007;
Matsumoto et al., 2007).
Figure 2: The molecular mechanism of the clock The transcription
factors CLOCK (CLK) and CYCLE (CYC) activate the transcription of
the target genes period (per), timeless (tim), vrille (vri) and Par
domain protein 1 (Pdp1) by binding to E boxes in their promotor
regions. The proteins regulate their own transcription and the
expression of other clock genes by either inhibiting CLK-CYC (first
feedback loop with PER and TIM) or by regulating clk transcription
(second feedback loop with VRI and PDP1). Cryptochrome (CRY)
mediates the light-induced degradation of TIM and is modulated by
Shaggy (SGG). Double-Time (DBT) leads to PER degradation if it is
not stabilized by TIM. Activators are colored in green, repressors
in red. For details see text. From Collins and Blau (2007)
Figure 3: The function of clockwork orange (cwo) CWO binds to
the E boxes of clock genes and represses their transcription.
Activators are colored in green, repressors in red. From Matsumoto
et al. (2007)
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Introduction
17
2.2 The clock network
In the brain of Drosophila melanogaster, about 150 neurons
express clock genes – they are
therefore called clock cells and comprise the circadian clock or
central pacemaker. According
to their anatomical location, the clock cells can be divided
into two main neuron clusters: the
dorsal neurons (DN) and the lateral neurons (LN). These clusters
can be further subdivided
into 3 DN clusters and 4 LN clusters. Each brain hemisphere
contains 16 DN1s, 2 DN2s and 40
DN3s, as well as 5 small ventral lateral neurons (sLNv), 4 large
ventral lateral neurons (lLNv),
6 dorsal lateral neurons (LNd) and 3 lateral posterior neurons
(LPN) (see Figure 4). Of the 5
sLNvs, only 4 express the neuropeptide pigment-dispersing factor
(PDF), as do the lLNvs, while
the 5th sLNv does not (Helfrich-Förster, 1995; Helfrich-Förster,
2007; Peschel and Helfrich-
Förster, 2011).
While all clock cells interact to produce normal rhythms of
behavior, the LNs seem to be the
most important pacemakers as they were shown to be both
necessary and sufficient for
normal locomotor activity rhythmicity (Ewer et al., 1992; Renn
et al., 1999). For eclosion
rhythmicity, the LNs are necessary, but not sufficient (Myers et
al., 2003).
Figure 4: The clock neurons in the adult brain of Drosophila
melanogaster The clock neurons can be divided into dorsal (DN) and
lateral (LN) neuron clusters. These clusters can be further
subdivided into DN1, DN2 and DN3 as well as small ventral lateral
neurons (sLNv), large ventral lateral neurons (lLNv), dorsal
lateral neurons (LNd) and lateral posterior neurons (LPN). From
Helfrich-Förster (2007)
-
Introduction
18
Other tissues outside of the brain were shown to express clock
genes that oscillate
autonomously (Plautz et al., 1997). As opposed to the central
clock in the brain, they are called
peripheral clocks and can be found in many tissues, for example
in the antennae (Plautz et al.,
1997), the Malpighian tubules (Giebultowicz and Hege, 1997), the
fat body (Xu et al., 2008)
and the prothoracic gland (PG) (Emery et al., 1997). They can be
independent of the central
clock or coupled to it, but can all be directly entrained by
light (reviewed in Ito and Tomioka,
2016).
2.3 Entrainment pathways of the circadian clock
2.3.1 Light entrainment
Drosophila has one internal and three external photoreceptive
systems: the blue light
receptor Cryptochrome (CRY) as internal photoreceptor and the
compound eyes, the ocelli
and the Hofbauer-Buchner-eyelets (HB-eyelets) as external
photoreceptive structures (Figure
4) (Rieger et al., 2003). The compound eye consist of about 800
ommatidia, each containing 8
photoreceptor cells, called R1 to R8. Each photoreceptor cell
expresses one type of rhodopsin
(Rh), the visual pigments of the eye. The outer photoreceptor
cells R1 to R6 express Rh1, while
the inner photoreceptor cells R7 and R8 express RH3 and Rh5 (in
the pale type) or Rh4 and
Rh6 (in the yellow type), respectively (reviewed in Montell,
2012). The HB-eyelet consists of 4
neurons between the retina and the lamina of the compound eyes
(Hofbauer and Buchner,
1989). It develops from the Bolwigs organ, which depicts the
photoreceptors in larvae. The
HB-eyelet expresses Rh6 and its projections innervate the LNvs
(Helfrich-Förster et al., 2002).
The ocelli are three simple eyes on the adult head (Goodman,
1970; Hu et al., 1978) that
express Rh2 (Pollock and Benzer, 1988). While all three
structures were shown to contribute
to locomotor rhythmicity under light entrainment, the compound
eyes are the main organs to
adapt the phase of activity and mediate the masking effect of
light (Rieger et al., 2003). Action
spectra for eclosion rhythmicity showed a peak of sensitivity
between 420 nm and 480 nm,
while wavelengths longer than 540 nm could not shift eclosion
any more (Frank and
Zimmerman, 1969).
CRY is sensitive to light in the UV-A/blue range (Van
Vickle-Chavez and Van Gelder, 2007) and
is expressed in all five sLNvs, the four lLNvs, in three of the
LNds and 8 DN1s, as well as non-
clock cells in the adult brain (Benito et al., 2008; Yoshii et
al., 2008). The function of CRY in
resetting the clock under light entrainment is described in
Chapter 2.1. Flies mutant for cry
can entrain normally to light cycles, but need longer to
re-entrain after phase shifts (Emery et
-
Introduction
19
al., 1998; Stanewsky et al., 1998) and stay rhythmic under
constant light (Emery et al., 2000a;
Dolezelova et al., 2007a). Overexpression of CRY in clock cells
increases their photosensitivity
(Emery et al., 2000b). For eclosion, contradictory results for
cry mutants have been reported:
while Myers et al. (2003) reported arrhythmic eclosion (Myers et
al., 2003), Mealey-Ferrara et
al. (2003) and Dolezelova et al. (2007a) found eclosion to stay
rhythmic (Mealey-Ferrara et al.,
2003; Dolezelova et al., 2007b), even under constant light
(Dolezelova et al., 2007a). In cry
mutants, the circadian oscillators in peripheral tissues are
largely arrhythmic (Krishnan et al.,
2001; Levine et al., 2002) and PER and TIM oscillations in the
PG exhibit smaller phase-
responses (Morioka et al., 2012).
2.3.2 Temperature entrainment
Temperature is a strong Zeitgeber for Drosophila, and an
amplitude of 2-3°C is enough to
synchronize locomotor activity behavior (Wheeler et al., 1993)
and eclosion (Zimmerman et
al., 1968b). Temperature cycles can entrain behavior together
with light (Yoshii et al., 2009)
or under constant darkness (Stanewsky et al., 1998; Yoshii et
al., 2002; Yoshii et al., 2005) and
even under constant light, that otherwise renders behavior
arrhythmic (Yoshii et al., 2002;
Glaser and Stanewsky, 2005; Yoshii et al., 2005). This
entrainment is clock-dependent, as per
mutants can merely react to temperature changes, but not entrain
any more (Wheeler et al.,
1993).
Many tissues and organs express per and tim (reviewed in Hall,
1995), and their oscillations
can be entrained to temperature, implying that temperature
entrainment is a cell-
autonomous mechanism (Glaser and Stanewsky, 2005). The existence
of a cell-autonomous
thermoreceptor, comparable to CRY for light detection, was
hypothesized (Glaser and
Stanewsky, 2005). However, Sehadova et al. (2009) reported that
isolated brains are able to
synchronize to light cycles, but not to temperature cycles. The
clock neurons are therefore
dependent on input from other neurons or tissues (Sehadova et
al., 2009).
Two mutant fly strains have been identified that are not able to
entrain to temperature cycles:
norpA and nocte (Glaser and Stanewsky, 2005). NorpA encodes for
phospholipase C, which
plays a role in the thermosensitive splicing of per as an
adaption to seasonal changes (Collins
et al., 2004; Majercak et al., 2004) and is also involved in the
phototransduction cascade of
the photoreceptors reviewed in Hardie (2001). Nocte seems to
have a specific function in the
entrainment to temperature cycles as flies mutant for nocte are
still able to entrain to light
cycles and their clocks are temperature compensated (Glaser and
Stanewsky, 2005).
-
Introduction
20
Knockdown of nocte in peripheral tissues impairs their
temperature synchronization and leads
to arrhythmic behavior (Sehadova et al., 2009). The chordotonal
organs (ch organs) were
identified to play a role in temperature entrainment, although
they possess no functional clock
(Sehadova et al., 2009). Nocte mutants show strongly deformed ch
organs which may cause
the observed impairments in temperature entrainment. Further
candidates for the
entrainment to thermocycles are the DN2 and LPN clock neurons
that showed stronger per
oscillations under temperature cycles than under light cycles
(Yoshii et al., 2005; Busza et al.,
2007; Miyasako et al., 2007; Picot et al., 2009).
2.4 The clock-related roles of pigment-dispersing factor
(PDF)
The neuropeptide pigment-dispersing factor (PDF) was first
isolated and characterized in
Drosophila in 1998 (Park and Hall, 1998) and soon identified as
the main output factor of the
clock cells in the brain (Renn et al., 1999; Helfrich-Förster et
al., 2000). Flies mutant for pdf or
with ablated PDF neurons show arrhythmic locomotor activity
(Renn et al., 1999) as well as
eclosion (Myers et al., 2003) under DD. The same is true when
the molecular clock is disrupted
in these neurons (Blanchardon et al., 2001) or when PDF is
overexpressed (Helfrich-Förster et
al., 2000). PDF is expressed in the four lLNvs and in four of
the five sLNvs (Helfrich-Förster,
1995), as well as in the tritocerebral PDF neurons that arise in
the mid-pupal stage and degrade
after eclosion (Helfrich-Förster, 1997). Besides PDF, the sLNvs
also express the small
Neuropeptide F (sNPF) (Johard et al., 2009). The receptor of
PDF, PDFR or Han, was discovered
in 2005 (Hyun et al., 2005; Lear et al., 2005; Mertens et al.,
2005) and was shown to be
expressed in CRY positive clock neurons and other cells outside
the clock network (Hyun et al.,
2005; Im and Taghert, 2010; Im et al., 2011). Like pdf mutants,
flies mutant for han become
arrhythmic under DD (Hyun et al., 2005; Lear et al., 2005;
Mertens et al., 2005).
Eclosion in Drosophila melanogaster
Like all arthropods, Drosophila melanogaster has to shed its
rigid exoskeleton to be able to
grow. This process is called molting. Drosophila has three
larval stages (L1 to L3), called instars.
Each developmental transition from one instar to the next is
accompanied by an ecdysis. At
the end of the third instar, the larvae start to wander out of
the food and pupariate. As a
holometabolous insect, Drosophila undergoes a complete
metamorphosis, and the last
ecdysis including the emergence of the adult fly is called
eclosion. This critical behavior is
-
Introduction
21
regulated by the interplay of the circadian clock, hormones and
peptides that all act together
to generate a precisely timed eclosion motor pattern (reviewed
in Ewer, 2007).
Although eclosion happens only once in the lifetime of a fly, a
clear rhythm in the eclosion of
mixed-age populations of flies can be observed. Eclosion
profiles show the number of flies
that eclosed within a certain timeframe over several days. As
the number of eclosed flies
changes from day to day and between experiments, the number of
flies that eclosed per hour
is usually normalized to the total number of flies that eclosed
per day, which allows for a
comparison of different days during one experiment and between
experiments. The
percentage of eclosed flies per hour is depicted as a bar. The
light or temperature regime,
under which the experiment was conducted, is shown in colored
rectangles above the eclosion
profile.
When pupae are raised under light:dark cycles (LD), most flies
eclose during the light phase,
and only few during the dark phase. The onset of light is
accompanied by a lights-on peak in
eclosion, which is a masking effect of the light: a startle
response induced directly by the
Zeitgeber, but not regulated by the circadian clock (Figure
5).
Figure 5: Eclosion rhythm under light:dark conditions (LD)
Eclosion profiles of the control strain w1118 under LD conditions.
Each bar represents the percentage of eclosed flies per hour
normalized to the number of eclosed flies per day. The black and
yellow rectangles on top represent the light regime (yellow: light
phase; black: dark phase).
-
Introduction
22
As eclosion is a circadian behavior, eclosion rhythms persist
also under constant darkness
conditions (DD). There are no distinct lights-on peaks any more,
and the eclosion events are
more broadly distributed. The beginning of the free-run after
around two days results in even
broader distributed eclosion events. As the free-running period
is usually slightly longer than
24 hours, the peaks are shifted a bit later each day. Under
constant conditions, the time is no
longer defined by the Zeitgeber, and is therefore called
circadian time (Figure 6).
Figure 6: Eclosion rhythm under constant darkness conditions
(DD) Eclosion profiles of the control strain w1118 under DD
conditions. Each bar represents the percentage of eclosed flies per
hour normalized to the number of eclosed flies per day. The black
rectangles on top represent the light regime (black: dark
phase).
The characterizing features of eclosion rhythms are the
amplitude, the period and the gate.
The amplitude describes the height of the eclosion peak, i.e.
how many flies eclosed during
one hour. The period is the time between two eclosion peaks.
Under LD conditions it will be
synchronized to the given Zeitgeber, while under DD conditions
it represents the endogenous
free-run period. The gate is the number of hours per day in
which eclosion occurs (Figure 7).
The gate is the “allowed zone” for eclosion, which is determined
by the circadian clock
(Pittendrigh, 1954). Flies can eclose only during this gate, and
if they reach the pharate state
too late at one day after the gate has closed, they have to wait
until the gate opens again the
next day to eclose (Pittendrigh and Skopik, 1970). Through
gating, eclosion is limited to a
specific time of the day.
-
Introduction
23
Figure 7: General features of eclosion rhythms The general
features that describe an eclosion profile are the amplitude, that
represents the number or percentage of flies that eclosed during
one hour; the period, that is defined as the time between two
consecutive eclosion peaks; and the gate, which gives the timeframe
in which eclosion happens during one day.
3.1 Hormonal regulation of eclosion
The ring gland (RG) of larvae depicts the most important
endocrine organ for the regulation
of development. It consists of three parts, the prothoracic
gland (PG), the corpora allata (CA)
and the corpora cardiaca (CC) and is located behind the brain in
close association with the
aorta (Figure 8). The two main antagonists regulating
developmental transitions are Juvenile
Hormone (JH), which is produced in the CA, and ecdysone (20E),
which is produced in the PG.
The production of 20E is induced by the prothoracicotropic
hormone (PTTH) via its receptor
Torso (for details see Chapter III). Each molt or developmental
transition is preceded by a pulse
of 20E. As long as JH titers are high, metamorphosis is
repressed and the molt ends in the next
larval stage. Decreasing JH titers allow 20E to induce the
wandering behavior, pupariation,
pupal metamorphosis and finally eclosion of the adult fly
(reviewed in Di Cara and King-Jones,
2013) (Figure 9). The titers of JH are bound to checkpoints
during the development: the
threshold size between second and third instar, as well as the
minimal viable weight and the
critical weight during the last larval stage. These checkpoints
ensure the larvae to survive the
metamorphosis into a fertile adult. When the checkpoint for
critical weight is passed, JH titers
decrease and metamorphosis is induced. In contrast, shortweight
provokes molting which
results in another larval stage (reviewed in Mirth and
Shingleton, 2012). The PG was identified
as the size-assessing tissue for these checkpoints, as changing
the size of the PG influenced
the size of the adult fly (Mirth et al., 2005). The PG was
reported to be mostly degenerated
shortly before eclosion (Dai and Gilbert, 1991), but newer
studies indicate that it degenerates
only after eclosion (Mareike Selcho, personal
communication).
-
Introduction
24
Figure 8: The ring gland (RG) of Drosophila melanogaster The RG
is located behind the larval brain and consists of the prothoracic
gland (PG; blue-green), the corpora allata (CA; purple) and the
corpora cardiaca (CC; yellow). The endings of the
prothoracicotropic hormone (PTTH) neurons (grey) terminate on the
PG portion of the RG.
Figure 9: Levels of hormones regulating eclosion during
development Each developmental transition is preceeded by a pulse
of ecdysone (20E; red line). As production and secretion of
ecdysone is induced by the prothoracicotropic hormone (PTTH; purple
line), PTTH peaks can be detected before 20E pulses. As long as
titers of the Juvenile Hormone (JH; green line) are elevated, the
20E pulses induce transition to another larval stage. When the
critical weight is reached, JH titers decrease and 20E induces
wandering behavior, pupariation and eclosion. From Lange et al.
(2010)
11 | P a g e
Introduction
Figure 4 - Hormone levels during development of insects: the
figure is an idealized illustration of hormone
fluctuations in insect larva and pupa and has been fitted to the
life cycle of D. melanogaster . Peaks in PTTH
concentrations give rise to an increase in the 20E
concentration. The JH titer declines in the end of the second
instar followed by small peaks in the PTTH titer in the
beginning of the third larval instar. The third instar
terminates with a large peak in PTTH, and subsequently the 20E
concentration in creases, permitting the larva
to go into the prepupal and pupal stage. Just before pupariation
the larva begins wandering behaviour to find
an appropriate location for pupariation. Critical weight is a
developmental checkpoint after which the process
of metamorphosis cannot be stopped. The terminal growth period
(TGP) is the period between obtainment of
critical weight and the cessation of growth. Based on (Edgar,
2006; Layalle et al., 2008; Nijhout, 2003;
Riddiford, 1993; Schubiger et al., 1998) .
-
Introduction
25
3.2 Circadian regulation of eclosion
The most important clock cells for rhythmic eclosion are the
PDF-expressing LNs:
overexpression of PDF or disturbance of the molecular clockwork
in the LNs as well as their
ablation results in arrhythmic eclosion under constant
conditions (Helfrich-Förster et al., 2000;
Blanchardon et al., 2001; Myers et al., 2003). Observation of
per and tim cycling revealed a
functional clock in the PG at the time of eclosion (Emery et
al., 1997; Myers et al., 2003) and
disruption of this clock leads to arrhythmic eclosion as well
(Myers et al., 2003). Therefore,
both the central clock in the brain and the peripheral clock in
the PG are required to maintain
eclosion rhythmicity. Morioka et al. (2012) reported that per
cycling is enhanced by signals
from the CNS and that light input to the PG is mediated through
the CNS (Morioka et al., 2012).
The interconnection of both clocks is unknown, but anatomical
studies suggest that the sLNvs
innervate the PG indirectly via the PTTH neurons whose endings
terminate on the PG
(Siegmund and Korge, 2001). It was shown in Manduca sexta,
Bombyx mori and Rhodnius
prolixus that PTTH is released only during a clock-regulated
gate and that the cycling in PTTH
release coincides with a cycling of 20E levels (Truman, 1972;
Truman and Riddiford, 1974;
Ampleford and Steel, 1985; Satake et al., 1998). Whether PTTH
and 20E are rhythmically
released in Drosophila is unknown. The titers of 20E are very
low at the end of pupal
development (Handler, 1982) (Figure 9), and so far it was not
possible to detect 20E synthesis
(Dai and Gilbert, 1991) or rhythmic release of 20E (Handler,
1982) on the day prior to eclosion.
Another factor influencing eclosion rhythmicity is the
RNA-binding protein LARK (Jackson,
1993; McNeil et al., 1998). LARK is expressed pan-neuronally and
affects eclosion rhythmicity
when overexpressed or knocked down in EH, CCAP or PDF neurons
and tim expressing cells
(Schroeder et al., 2003; Sundram et al., 2012). Neither cell
morphology nor clock protein
cycling are affected, but PDF neurons show decreased PDF
immunoreactivity in lark mutants.
Therefore, a function in the output of clock cells is suggested
(Schroeder et al., 2003).
Knockdown of lark in the PG also leads to arrhythmic eclosion
(Sundram et al., 2012).
3.3 Peptidergic regulation of eclosion in Drosophila
melanogaster
Each ecdysis is a sequence of specific motor patterns, divided
into pre-ecdysis, ecdysis and
post-ecdysis behaviors. Eclosion is orchestrated by a signaling
cascade of peptides that
ensures the right succession of each of these behaviors. The
main regulators are ecdysis-
triggering hormone (ETH), which is secreted by peripheral
epitracheal cells, also called Inka
-
Introduction
26
cells, eclosion hormone (EH) from the Vm neurons in the brain
and crustacean cardioactive
peptide (CCAP) from CCAP neurons in the brain and thoracic
ganglion. ETH and EH form a
positive feedback loop, where each peptide induces the release
of the other until all stores
are depleted (Ewer et al., 1997). Eclosion starts with a release
of ETH from the Inka cells (Figure
10 ①) (Zitnan et al., 1996; Ewer et al., 1997) which induces via
its receptor the release of EH
from the Vm neurons (Figure 10 ②) (Ewer et al., 1997; Kim et
al., 2006b). EH then leads to
the rapid secretion of ETH from the Inka cells via its receptor
(Chang et al., 2009) and an
increase of cGMP levels (Figure 10 ③) (Clark et al., 2004).
Studies in eh null-mutants showed
that ETH is secreted normally even without the EH cells and that
the first release of ETH was
not accompanied by an increase of cGMP levels, suggesting that
the initial release of ETH is
independent of EH (Clark et al., 2004). In Manduca sexta,
Corazonin (Crz) was shown to trigger
this first release of ETH from Inka cells (Kim et al., 2004)
while no comparable starting signal
is known in Drosophila.
Mainly ETH activates further target neurons via its receptor
(Kim et al., 2006a) and thereby
promotes together with EH pre-ecdysis (Figure 10 ④) and ecdysis
behavior (Figure 10 ⑤).
Among the main targets of ETH and EH are the CCAP neurons
(McNabb et al., 1997a; Kim et
al., 2006b). CCAP inhibits pre-ecdysis behavior and further
promotes ecdysis behavior (Figure
10 ⑥) (Gammie and Truman, 1997). CCAP neurons express
additionally to CCAP also
myoinhibitory peptide (MIP) and bursicon (Kim et al., 2006a;
Vömel and Wegener, 2007).
While MIP acts together with CCAP to promote ecdysis behavior,
bursicon is necessary for
post-ecdysis behaviors like sclerotonization, pigmentation and
wing inflation (Figure 10 ⑦)
(Dewey et al., 2004; Luo et al., 2005; Peabody et al., 2008;
Lahr et al., 2012; Kim et al., 2015;
Krüger et al., 2015).
Ecdysis must start only at the end of the molt when the old
cuticle can be shed and the new
cuticle is already synthesized (Ewer, 2007). A main regulator of
this timing is 20E, as the
increase of 20E levels induces the production of ETH (Zitnan et
al., 1999) via an Ecdysone-
response element (Park et al., 1999), raises the sensitivity of
the CNS to ETH (Zitnan et al.,
1999) and regulates the secretory competence of the Inka cells
(Kingan and Adams, 2000; Cho
et al., 2013). The system is now “armed” and injection of ETH
(Zitnan et al., 1996) or EH
(Truman et al., 1983) at this stage can induce premature
eclosion. But Inka cells become
sensitive to EH only after the drop of 20E levels, preventing
the positive feedback loop from
starting too early (Ewer et al., 1997; Zitnan et al., 1999).
-
Introduction
27
Figure 10: The peptide signaling cascade regulating eclosion
behavior Eclosion starts with an initial release of the
ecdysis-triggering hormone (ETH) from the epitracheal Inka cells ①
which induces release of the eclosion hormone (EH) from the Vm
neurons with somata in the brain ②. EH and ETH form a positive
feedback loop and EH promotes complete depletion of ETH ③. EH and
ETH activate target neurons and thereby promote pre-ecdysis ④ and
ecdysis behavior ⑤. Among their main targets are crustacean
cardioactive peptide (CCAP) neurons that also express the peptides
myoinhibitory peptide (MIP) and bursicon. CCAP and MIP inhibit
pre-ecdysis behavior and further promote ecdysis behavior ⑥.
Bursicon is necessary for post-ecdysis behaviors like hardening and
pigmentation of the cuticle and wing inflation ⑦. After Ewer
(2005).
Aim of the present dissertation research
The general aim of this dissertation research is to dissect the
connection between the central
clock and the peptides regulating eclosion behavior on different
organization levels.
The first part focuses on the impact of Zeitgebers and PDF
signaling on natural eclosion
rhythmicity. To this end, a new eclosion monitoring system was
developed to allow the study
of eclosion behavior under natural conditions. For the analysis
of the complex interactions
between clock and environment, a statistical model was
established in a collaborative effort
within the collaborative research center (CRC) 1047 “Insect
timing”.
The second part of the present work focuses on the interaction
of the central clock and the
peripheral clock in the PG (Emery et al., 1997), as both systems
were shown to be necessary
for rhythmic eclosion (Myers et al., 2003). The PTTH signaling
cascade seemed to be a good
candidate pathway for this connection, as PTTH neurons terminate
on the PG (Siegmund and
Korge, 2001) and their dendrites are in close proximity of
arborizations from PDF neurons
(McBrayer et al., 2007; Selcho et al., unpublished-a). Indeed,
first results from our group
implied their role in eclosion rhythmicity (Chen, 2012).
Ablation and silencing experiments
were conducted under light and temperature entrainment, and
possible cycling of Ptth mRNA
levels was analyzed by means of qPCR. To dissect the connection
between the PDF and PTTH
neurons, the receptors of the two peptides expressed in PDF
neurons, PDF (Helfrich-Förster,
1995) and sNPF (Johard et al., 2009), were knocked down in PTTH
neurons. The receptor of
-
Introduction
28
PTTH, Torso, was knocked down pan-neuronally and in the PG and
its expression pattern was
analyzed by PCR.
The third part of this dissertation research focuses on the
connection between the central
clock and the peptide cascade regulating eclosion behavior. CCAP
is a main promotor of
ecdysis behavior (Gammie and Truman, 1997; McNabb et al., 1997a)
and the synaptic endings
of CCAP-expressing neurons overlap with clock neurons and the
PDFTri neurons (Park et al.,
2003) for which a potential role in the circadian timing of
eclosion is hypothesized (Helfrich-
Förster, 1997). Ablation of CCAP neurons was shown to impair
eclosion rhythmicity (Park et
al., 2003) and in a first set these experiments were repeated.
During this thesis, a null-
mutation for ccap was published which showed that CCAP does not
affect eclosion rhythmicity
(Lahr et al., 2012). This mutant was tested in the present work
under light and temperature
entrainment as well as under natural conditions for possible
consequences on eclosion
rhythmicity.
The fourth part of this thesis was designated to identify
peptidergic starting signals for
eclosion behavior, comparable to the role of Corazonin in
Manduca (Kim et al., 2004). An
optogenetic screen was established in which specific peptidergic
neurons were activated to
reveal their potential to trigger eclosion.
-
Material & Methods
29
Materials and Methods
Eclosion assays
All flies were raised on standard Drosophila food medium (0.8%
agar, 2.2% sugarbeet syrup,
8.0% malt extract, 1.8% yeast, 1.0% soy flour, 8.0% corn flour
and 0.3% hydroxybenzoic acid).
For each eclosion experiment, at least 3 culture vials with a
volume of 165 ml (K-TK; Retzstadt,
Germany) were prepared with a minimum of 50 flies each. Flies
were transferred to new
culture vials every 2 to 3 days and were entrained either under
LD (see 1.1) or WC conditions
(see 1.2). For each experiment, pupae with an age span of one
week were used. For
experiments under natural conditions, at least 6 culture vials
with a volume of 165 ml (K-TK;
Retzstadt, Germany) were prepared and flies were transferred to
new culture vials twice a
week. New adult flies were added regularly to compensate loss of
old flies.
1.1 Würzburg Eclosion Monitor (WEclMon)
The eclosion monitors were custom-made by the Biocenter workshop
(Johann Kaderschabeck)
and the department’s electronician Konrad Öchsner. First,
eclosion plates were made out of
acrylic glass plates with 1,000 2 mm high platforms and a
built-in metal frame to reduce static
current. Illumination came from below by lighting plates
consisting of LED Stripes in red (12V
SMD 3528 Red 60 LED/m; λ=635 nm), white (12V SMD 3528 Cool White
60 Led/m; max.
emission at λ=450 nm) or infrared (YB-G3528IR60F08N12, IR850,
12V; λ=850 nm) fixed around
an acrylic glass plate by a metal frame. For the activation of
channelrhodopsin, blue LEDs
(LXHL-LB5C 470 nm) were additionally built into the
monitors.
As recording cameras, either a Logitech HD - C920 (Logitech,
Romanel-sur-Morges,
Switzerland) or the board camera Delock 95955 (Tragant Handels-
und Beteiligungs GmbH,
Berlin, Germany) were used. Camera filters to eliminate normal
day light were either 625/30
ET bandpass filters (AHF analysentechnik AG, Tübingen, Germany)
for the red LED light with a
wavelength of λ=635 nm, or unexposed yet developed photographic
film for infrared light. For
intervalled image recording the freely available software Yawcam
(http://www.yawcam.com)
was used. Image and data analysis was performed in FiJi
(Schindelin et al, 2012) using the
“Eclosion bar” macro developed by Martin Fraunholz, Department
of Microbiology, University
of Würzburg, described in Chapter I.
http://www.yawcam.com/
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Material & Methods
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1.2 Eclosion assays under light entrainment
Flies were raised under a light regime of 12 hours light and 12
hours darkness, with 20°C and
65% relative humidity. 12 to 17 days old pupae were collected
and transferred into the
monitoring systems. Eclosion was monitored using either the
Trikinetics (TriKinetics,
Massachusetts, USA) or the WEclMon monitoring systems (for
description see Chapter I and
Ruf et al, in preparation).
For experiments using the TriKinetics monitoring system, pupae
attached to the walls of the
culture vial were rinsed with water and carefully removed with a
spatula. The collected pupae
were washed in water to remove rests of the food medium and then
separated on filter paper
(Whatman® Blotting Papers). After drying, pupae were fixed on
plastic eclosion plates using a
thinly spread out cellulose-based glue (Auro Tapetenkleister Nr.
389; 1:30). At the end of the
light phase, the eclosion plates were mounted on top of glass
funnels within the monitors (day
0) and eclosion was monitored for one week at 20°C, either under
12 hours light and 12 hours
darkness (LD) or under constant darkness (DD). In the
TriKinetics monitoring system, freshly
eclosed flies are shaken from the eclosion plate by a tapping
solenoid that pushes down the
plate and the funnel once every minute. The freshly eclosed
flies with folded wings fall through
the funnel into beakers filled with water and detergent. While
falling through the funnel, the
flies interrupt an infrared beam, which is automatically counted
by the DAMSystem Collection
Software (DAMSystem303) and read out as a text file by the
recording computer.
For experiments using the WEclMon monitoring systems, pupae were
individually taken out
of the culture vial and glued onto a platform on the eclosion
plates by a drop of cellulose-
based glue (Auro Tapetenkleister Nr. 389; 1:30). At the end of
the light phase of day 0, the
eclosion plates were mounted in the eclosion monitors and
eclosion was monitored for one
week at 20°C, either under 12 hours light and 12 hours darkness
(LD) or under constant
darkness (DD). Infrared light (λ=850 nm) was given throughout
the experiment.
1.3 Eclosion assays under temperature entrainment
Flies were raised under constant red light (λ=635 nm) and 65%
relative humidity in climate
chambers (Plant Growth Chamber, DR-36NL, Percival Scientific,
Inc., Perry, USA) and
entrained to a temperature regime of 12 hours at 25°C and 12
hours at 16°C. The temperature
increase and decrease was ramped by 1°C every 10 minutes.
Eclosion was monitored using
the WEclMon monitoring systems. After 13 to 18 days, the pupae
were individually mounted
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Material & Methods
31
on eclosion plates and placed in the monitors. Eclosion was
monitored for one week either
under 12:12 hours 25°C:16°C (WC) or under a constant temperature
of 20°C (CC) as well as
65% relative humidity and constant red light (λ=635 nm) in the
same incubators.
1.4 Eclosion assays after optogenetic activation via
channelrhodopsin
Flies were raised under constant red light (λ=635 nm) and 65%
relative humidity in climate
chambers (Plant Growth Chamber, DR-36NL, Percival Scientific,
Inc., Perry, USA) and
entrained to a temperature regime of 12 hours at 25°C and 12
hours at 16°C. The temperature
increase and decrease was ramped by 1°C every 10 minutes.
Eclosion was monitored using
the WEclMon monitoring systems. After 14 to 19 days, the pupae
were individually mounted
on eclosion plates and placed in the monitors. Eclosion was
monitored for one week under
12:12 hours 25°C:16°C at 65% relative humidity and constant red
light (λ=635 nm) in the same
incubators. At day one of the experiment, blue light of 586
µW/cm2 (λ=470 nm) was given for
one hour at the indicated time point.
1.5 Eclosion assays under natural condition
The experiments under natural conditions were conducted from
July to October 2014 in an
enclosure at the bee station of the University of Würzburg
(Figure S 5). The enclosure was
shaded from direct sunlight by bushes and a dark tarpaulin. The
walls were lined with insect
mesh to allow air circulation and to reduce effects on the
relative humidity. The mesh also
kept off other insects and birds and, most of all, kept the
eclosed flies inside the enclosure
where they were caught with sticking paper and vinegar traps.
During their whole
development, flies were raised in culture vials in the enclosure
and prepared for eclosion
monitoring once pharate pupae had appeared. Eclosion was
monitored using the WEclMon
monitoring systems. At the day of preparation, the vials were
taken into the lab and pupae
were individually transferred onto the platforms of the eclosion
plates. On the same day, the
ready eclosion plates were then transferred back into the
enclosure and mounted in the open
WEclMon. Eclosion was monitored for one week under constant red
light (λ=850 nm) without
further manipulating the pupae. In parallel, temperature, light
intensity and relative humidity
were measured directly in the eclosion monitors using a
datalogger (MSR 145S, MSR
Electronics GmbH Seuzach, Switzerland).
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Material & Methods
32
1.6 Analysis of rhythmicity
Rhythmicity and period length of the eclosion profiles were
analyzed using MATLAB
(MathWorks, Inc., Natick, USA) and the appropriate Matlab
toolbox (Levine et al., 2002).
Statistical analysis with one-way ANOVA followed by Tukey
post-hoc test and independent-
samples t-test were performed with IBM® SPSS® Statistics
software (version 20).Graphical
output
Graphs of eclosion profiles were compiled in R (version 3.2.0;
The R Project for Statistical
Computing).
Molecular analysis
2.1 Analysis of torso expression via Reverse Transcription
Polymerase-Chain-Reaction
(RT-PCR)
Isolation of total RNA
For total RNA extraction, the Quick-RNA™ MicroPrep Kit from Zymo
Research (Irvine, USA)
was used and all steps were performed according to the
manufacturer´s instruction.
Pharate and adult Canton S flies not older than 3 days were
dissected in HL3.1 medium (Yanfei
Feng, 2004). Central brains, optic lobes and retinae of 15 flies
of each group were transferred
into 300 µl RNA lysis buffer on ice. Additionally, the gonads of
10 adult males and females, as
well as the complete abdomen of 10 male and female pharate
flies, were collected in 300 µl
RNA lysis buffer on ice. As positive control, brains with ring
glands from Canton S L3 wanderer
larvae were collected and treated the same way. The tissues were
mechanically homogenized
by means of a plastic pestle. Following washing and
centrifugation steps as described in the
manufacturer´s manual, total RNA was eluted in 8 µl water.
Reverse Transcription (RT)
For cDNA synthesis, the QuantiTect® Reverse Transcription Kit
from Qiagen (Venlo,
Netherlands) was used and all steps were performed according to
the protocol provided by
the manufacturer.
Genomic DNA remnants were removed by adding 1 µl of gDNA wipeout
buffer to 6 μl of the
eluted RNA. Following incubation at 42°C for 2 minutes, the
samples were placed for 2 minutes
at 4°C to suppress the reaction. 3 µl mastermix composed of 2 μl
RT Buffer, 0.5 μl RT Primer
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Material & Methods
33
Mix and 0.5 μl Reverse Transcriptase were added to the RNA and
the samples were placed
into a thermocycler (MJ Mini, Bio-Rad; Hercules, USA) to perform
the following temperature
steps: 30 minutes at 42°C, 3 minutes at 95°C and 2 minutes at
4°C. Finally, 40 µl water were
added and the cDNA samples were frozen and stored at -20°C.
PCR
To validate the expression of torso mRNA in the isolated fly
tissues, the reverse-transcribed
cDNA was used in a PCR assay. Primers were designed by means of
the Primer-BLAST function
of NCBI (http://www.ncbi.nlm.nih.gov/tools/primer-blast/) and
synthesized by Sigma-Aldrich
(Munich, Germany). As a positive control, expression of
α-tubulin was measured in the same
samples. The primer sequences are provided in Table 1.
Composition of the PCR reaction mix
is presented in Table 2, the PCR temperature program in Table 3.
PCRs were performed with
a MJ Mini thermocycler (Bio-Rad, Hercules, USA). As negative
control, the cDNA was
substituted by pure water. The PCR products were separated by
agarose gel electrophoresis
on a 1% gel (Roti®garose, Agarose Standard; Carl Roth,
Karlsruhe, Germany) with the
GeneRuler® 100 bp Plus DNA ladder from Fermentas (now
ThermoFischer Scientific; Waltham,
USA) in a Mini-Sub® Cell GT system from Bio-Rad (Hercules, USA)
and documented using the
E‐Box version 15.05 (Vilber Lourmat; Eberhardzell, Germany) and
evaluated with E‐Capt
version 15.06.
Table 1: Primers used for the analysis of torso expression
Target Primer name Primer sequence (5’-3’) Product size
torso torso 7 forward TCATCGAGAGGGCAACATGG’ 667 bp
torso 7 reverse CACAGTGGACAGCATCGAGT
α-tubulin tubulin forward TCTGCGATTCGATGGTGCCCTTAAC 198 bp
tubulin reverse GGATCGCACTTGACCATCTGGTTGGC
http://www.ncbi.nlm.nih.gov/tools/primer-blast/
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Material & Methods
34
Table 2: PCR reaction
JumpStart™ REDTaq® ReadyMix™ Reaction Mix (Sigma-Aldrich;
Missouri, USA) 12.5 µl
forward primer 10 µM 1 µl
reverse primer 10 µM 1 µl
cDNA template 5 µl
H2O 5.5 µl
total reaction volume 25 µl
Table 3: PCR program
2.2 Analysis of the temporal expression of Ptth mRNA via
quantitative Real-time
Polymerase-Chain-Reaction (qPCR)
mRNA extraction
For mRNA extraction, the Quick-RNA™ MicroPrep Kit from Zymo
Research (Irvine, USA) was
used and all steps were performed according to the
manufacturer´s protocol.
Canton S flies were reared under a light regime of 12 hours
light and 12 hours darkness at a
constant temperature of 20°C and 65% relative humidity in big
culture vials. Half of the vials
were transferred into DD conditions at the end of the light
phase after 11 days.
After 17 days, sample collection started. Heads of 20 pharate
flies per light regime were
collected every 4 hours (ZT 0, ZT 4, ZT 8, ZT 12, ZT 16, ZT 20)
in 300 µl RNA lysis buffer and
immediately frozen in liquid nitrogen. For the time points at
night or under DD, flies were
95°C 5 minutes
95°C 30 seconds
63°C 30 seconds
72°C 1 minute
72°C 5 minute
4°C hold
35 cycles
https://de.wikipedia.org/wiki/Missouri
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Material & Methods
35
prepared under red light of λ=635 nm. For each time point, two
samples were collected on 5
consecutive days. The tissues were manually homogenized with a
plastic pestle. Following the
washing and centrifugation steps described in the manufacturer´s
protocol, mRNA was eluted
in 8 µl water and the mRNA samples were frozen and stored at
-80°C.
Reverse Transcription (RT)
For cDNA synthesis the QuantiTect® Reverse Transcription Kit
from Qiagen (Venlo,
Netherlands) was used and all steps were performed according to
the manufacturer´s
protocol.
To make sure that every sample contains the same amount of cDNA,
the amount of mRNA in
each sample was measured by a NanoDrop™ 2000c spectral
photometer from ThermoFischer
Scientific (Waltham, USA). For each reaction, 1 µg mRNA, diluted
in pure water, was used and
reverse-transcribed into cDNA. 1 µl gDNA wipeout buffer was
added according to the volume
of diluted RNA and incubated for 2 minutes at 42°C until the
reaction was suppressed by
placing the samples for 2 minutes at 4°C. 3 µl mastermix
composed of 2μl RT buffer, 0.5μl RT
primer mix and 0.5μl reverse transcriptase were added to the
mRNA and the samples were
placed into a MJ Mini Personal Thermal Cycler from Bio-Rad
(Hercules, USA) to perform the
following temperature steps: 30 minutes at 42°C, 3 minutes at
95°C and 2 minutes at 4°C.
Finally, 40 µl water were added and the cDNA samples were frozen
and stored at -20°C.
qPCR
For quantification of Ptth mRNA expression, qPCR was performed
with the previously
obtained cDNA. Specific primers were designed using the
Primer-BLAST function of NCBI
(http://www.ncbi.nlm.nih.gov/tools/primer-blast/) and
synthesized by Sigma-Aldrich
(Missouri, USA). α-tubulin and RpL32 were chosen as reference
genes for later normalization
(Ponton et al., 2011). Primer efficiency was tested by a
dilution series (1:10³ to 1:106) utilizing
the PCR product from a standard PCR for each primer tested. The
sequences for the primers
used are shown in Table 4. The optimal annealing temperature of
60.5°C for all three primers
was determined through a gradient PCR with temperatures ranging
from 59°C to 67°C. The
ideal cDNA concentration of 100 ng per sample was identified by
testing different
concentrations between 10 ng and 100 ng. 5 biological replicates
were measured in three
technical replicates. Additionally, non-template control
reactions, for which cDNA was
http://www.ncbi.nlm.nih.gov/tools/primer-blast/https://de.wikipedia.org/wiki/Missouri
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Material & Methods
36
substituted by water, served as negative controls. qPCR
reactions were mixed in Eppendorf®
twin.tec PCR 96 well plates and sealed with adhesive Eppendorf®
Masterclear real-time PCR
film and the qPCR was performed in an Eppendorf Mastercycler
(Hamburg, Germany) using
the SensiMix™ SYBR® No-ROX Kit from Bioline (London, UK). The
qPCR reaction components
are shown in Table 5, the qPCR program in
Table 6.
Table 4: Primers for qPCR
Target Primer name Primer sequence (5’-3’) Product
size
Tm
Ptth ptth 7
forward
AAAGGTAATCCGAGAGGCGG 136 bp 59.54°C
ptth 7
reverse
ATAATGGAAATGGGCAACCACG 59.57°C
α-tubulin tubulin 2
forward
TTTACGTTTGTCAAGCCTCATAG 149 bp 57.14°C
tubulin 2
reverse
AGATACATTCACGCATATTGAGTTT 57.21°C
RpL32 rpl32 2
forward
ATGCATTAGTGGGACACCTTCTT 130 bp 59.99°C
rpl32 2
reverse
GCCATTTGTGCGACAGCTTAG 60.47°C
Table 5: qPCR reaction
SensiMix SYBR® No-ROX (Bioline) 10 µl
forward Primer 10 µM 0.8 µl
reverse Primer 10 µM 0.8 µl
cDNA template (for a concentration of 20 ng/µl) 5 µl
H2O 3.4 µl
total reaction volume 20 µl
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Material & Methods
37
Table 6: qPCR program
qPCR data analysis
The SYBR-Green fluorescence signal was monitored in real time as
amplification curves. A
threshold for the fluorescence signal was automatically set by
the Eppendorf® Mastercycler
ep realplex software version 2.2 (Eppendorf®; Hamburg, Germany)
with consideration of the
noise band calculation. The number of PCR cycles until the
signal surpassed the threshold was
determined for each sample (cycle threshold, CT). The mean CT
value of the technical
triplicates was calculated. If values differed from the other CT
values by > 0.5 CT, they were
considered as outlier and excluded from data analysis. As the
difference between the
reference genes was rather large, calculations were performed
separately for each reference
gene. For each sample, the ΔCT value was calculated by
subtracting the CT value of the target
gene (Ptth) from one of the reference genes. The 2- ΔΔCT value
was calculated by subtracting
the median of the ΔCT values for each reference gene and light
regime from the ΔCT values
(ΔΔCT) and building the negative quadratic term of the ΔΔCT. The
mean of the 2- ΔΔCT values for
each tested time point was plotted with standard deviations. For
statistical analysis, a one-
way ANOVA was performed between the 2-ΔΔCT values for each time
point and reference gene
for LD and DD conditions. Statistics were performed with IBM®
SPSS® Statistics software
(Version 20).
95°C 2 minutes
95°C 5 seconds
60.5°C 10 seconds
72°C 15 seconds
melting curve
4°C hold
35 cycles
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Material & Methods
38
Fly strains
For targeted expression of transgenes with high spatial control,
the GAL4-UAS system was
employed (Brand and Perrimon, 1993). This binary system consists
of the yeast GAL4
transcriptional activator and its target sequence UAS (Upstream
Activation Sequence). Two
lines of transgenic flies are created: one comprising the GAL4
cloned downstream of a
particular enhancer sequence, and a second including the UAS
upstream of an effector gene.
By combination of these transgenic flies, effector genes are
expressed with a high spatial
control in the F1 generation (Brand and Perrimon, 1993). One of
the numerous extensions of
the GAL-UAS system that was used in this thesis is the selective
blocking of the GAL4 via the
GAL80 repressor (Suster et al., 2004). The fly strains used in
this thesis are summarized in
Table 7.
Table 7: Fly strains used in this thesis
Fly strain Reference
akh-GAL4 Lee and Park (2004), kind gift of Jae Park
w; ap/cyo-GAL4 Bloomington Drosophila Stock Center #3041
AstA-GAL4 Hergarden et al. (2012), kind gift of David
Anderson
Canton S Stern and Schaeffer (1943), stock collection
capa-GAL4 Bloomington Drosophila Stock Center #51969
Bloomington Drosophila Stock Center #51970
CCAP exc7 Lahr et al. (2012), kind gift of John Ewer
ccap-GAL4 Lahr et al. (2012), kind gift of John Ewer
w; UAS ChR2-XXL Dawydow et al. (2014), kind gift of Robert
Kittel
dh31-GAL4 Bloomington Drosophila Stock Center #51988
Bloomington Drosophila Stock Center #51989
dh44-GAL4 Bloomington Drosophila Stock Center #39347
https://en.wikipedia.org/wiki/Upstream_Activation_Sequence
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Material & Methods
39
Vienna Drosophila Resource Center VT039046
UAS dicer2 (II) Vienna Drosophila Resource Cente #24666
UAS dicer2 (III) Vienna Drosophila Resource Cente #24667
dms-GAL4 8 Sellami et al. (2012), kind gift of Jan Veenstra
yw;+;UAS dORKΔ-C1 Nitabach et al. (2002),
Bloomington Drosophila Stock Center #6586
yw;+;UAS dORKΔ-NC1 Nitabach et al. (2002),
Bloomington Drosophila Stock Center #6587
Eh-Gal4 McNabb et al. (1997a), kind gift of John Ewer
elav-GAL4; UAS dicer Bloomington Drosophila Stock Center
#25750
w; eth-Gal4 Park et al. (2002) , kind gift of Michael Adams
w; UAS grim;+ Wing et al. (1998),
Bloomington Drosophila Stock Center #9923
w+; han5304 Hyun et al. (2005), stock collection
hug-GAL4 Bader et al. (2007), kind gift of Michael Pankratz
Mai 316-Gal4 Siegmund and Korge (2001), kind gift of
Thomas Siegmund
mip-GAL4 mip-Gal4-4M (III), stock collection
npf-GAL4 Bloomington Drosophila Stock Center #25683
nSybGAL80 Harris et al. (2015)
nSybGAL80; mip-GAL4 crossed for this thesis
pdf-GAL4 Renn et al. (1999), kind gift of Paul Taghert
w+; pdf01 Renn et al. (1999), kind gift of Paul Taghert
UAS pdfr RNAi Vienna Drosophila Resource Center #42724
http://flybase.org/cgi-bin/uniq.html?FBst0006586%3Efbsthttp://flybase.org/cgi-bin/uniq.html?FBst0006586%3Efbsthttp://stockcenter.vdrc.at/control/product/~VIEW_INDEX=0/~VIEW_SIZE=100/~product_id=42724
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Material & Methods
40
w+; per01 Konopka and Benzer (1971), stock collection
w;+;phm-Gal4,UAS-gfp/TM6B Mirth et al. (2005), kind gift of
Naoki Yamanaka
ptth-Gal4 McBrayer et al. (2007), kind gift of Naoki
Yamanaka
snpf-GAL4 Bloomington Drosophila Stock Center #202196
w;; UAS snpfr RNAi Bloomington Drosophila Stock Center
#27507
tdc-GAL4 Cole et al. (2005)
th-GAL4 Friggi-Grelin et al. (2003)
trh-GAL4 Sitaraman et al. (2012)
w1118 stock collection
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Chapter I
41
Chapter I. Development of a new eclosion monitoring system
(WEclMon)
Introduction
Eclosion, the emergence of the adult fly from the puparium, is a
circadian behavior under
regulation of peptides in Drosophila. It was among the first
rhythmic animal behaviors
discovered (Bremer, 1926; Bünning, 1935; Kalmus, 1935; Kalmus,
1938; Kalmus, 1940). Later
on, chronobiologists like Pittendrigh, Bruce or Zimmermann used
eclosion in Drosophila as
model behavior to study the properties of circadian clocks
(Pittendrigh, 1954; Pittendrigh and
Bruce, 1959b; Skopik and Pittendrigh, 1967; Zimmerman et al.,
1968a) and eclosion assays
were also used to identify the first clock gene period (Konopka
and Benzer, 1971). Eclosion is
an ideal behavioral model to study the coherencies of the
internal clock and peptidergic
regulatory system as it is not influenced by the motivational
state of the animal, or by hunger,
aggression, social contacts or reproductive state. As eclosion
happens only once in the lifetime
of the fly, rhythmicity can only be observed when monitoring in
a bigger population of flies of
mixed-age.
Over time, different methods have been established to monitor
eclosion rhythms. The
simplest way is to empty the culture vials in defined intervals
and count the number of eclosed
flies per hand. Early systems developed were the so-called
bang-boxes, where the pupae were
glued to metal plates that where mechanically lifted and let
fallen down, so that freshly
eclosed flies were forced to fall through a funnel into
water-filled containers (Engelmann,
2003). These containers were automatically exchanged by a
turning device. The U.S. company
TriKinetics Inc. further developed and refined this system into
the commercially available
Drosophila Eclosion Monitor (http://www.trikinetics.com/). Here,
the pupae are glued onto
plastic discs which are placed upside-down (pupae facing
downwards) on top of a funnel. A
small weight periodically “hammers” onto the plastic disc and by
that shakes down freshly
eclosed flies in fixed time intervals, which fall through a
funnel and are automatically counted
by crossing an infrared-beam. Although very easy to handle, this
system has some
disadvantages. For example, the funnels are often blocked, so
that experiments cannot be
used for analysis. Also it is unclear whether the constant
tapping of the pupae has an influence
on the eclosion rhythm of the flies. As the TriKinetics monitors
are closed systems, they are
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Chapter I
42
not suited to monitor eclosion rhythms under natural conditions,
since temperature, humidity
and air cannot be freely exchanged.
Therefore a novel open monitoring system was developed that
would allow the pupae to
come in direct contact with light, temperature and humidity and
exclude other experimental
influence.
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Chapter I
43
Results
2.1 WEclMon
The Würzburg Eclosion Monitor (WEclMon) is a camera-based, open
system. This system
provides the advantages that pupae can come in direct contact
with abiotic environmental
factors, like light, temperature and relative humidity.
Moreover, no mechanical manipulations
are needed as in the Trikinetics Monitors. The WEclMon consists
of three parts: a digital
camera (Figure 11, a), an eclosion plate (Figure 11, b) and one
or two light plates (Figure 11,
c, c’).
The camera (Figure 11, a) is positioned centrally above the
eclosion plate. Essentially any
camera can be used, according to the requirements of the
experiment. A simple webcam
(Logitech C910) was chosen whose resolution was sufficient to
reliably and simultaneously
monitor eclosion events of a high number of pupae. To take
images in certain time intervals,
the freely available software Yawcam (http://www.yawcam.com/)
was used.
The eclosion plate (Figure 11, b) is an acrylic glass plate with
1,000 2 mm high platforms.
Platforms instead of notches were chosen, as otherwise flies
often were unable to leave their
puparium due to space restrictions. On each platform, one single
pupa is placed and fixed with
cellulose-based glue, which allows to monitor the eclosion
events of 1,000 flies in one
experiment.
Illumination comes from the light plates (Figure 11, c, c’),
which are positioned below the
eclosion plate. Each light plate consists of a LED stripe of a
defined wavelength fixed around
an acrylic glass plate and a metallic frame to reduce the static
current. To be able to monitor
eclosion during the night or during the dark phase,
respectively, constant red light of a long
wavelength is given constantly. In the beginning, a wavelength
of λ=635 nm was chosen that
flies should not be able to perceive (Salcedo et al, 1999) and
should not influence their
eclosion behavior (Frank and Zimmerman, 1969; Helfrich-Förster,
2002). First results however
showed a strong influence on eclosion rhythms after LD
entrainment (Figure S 1, Figure S 2).
In a new set of monitors, the use of infrared light with a
wavelength of λ=850 nm was
therefore established. Changes in light intensity, for example
at sunrise or sunset or the switch
on and off of light, impede the data analysis and give false
positive results. Therefore, optic
filters optimized for the applied illumination wavelength were
added to reduce these
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Chapter I
44
differences in illumination. These were either bandpass filters
for the wavelength λ=635 nm
or unexposed, yet developed photographic film for infrared
light.
All parts of the monitor are assembled on a wooden base and can
be covered by a metallic
light-tight case, so that each monitor is an independent unit
and many can be used in parallel
for different experiments.
Figure 11: The Würzburg Eclosion Monitor (WEclMon) Schematic
drawing (A) and picture (B) of the Würzburg Eclosion Monitor
(WEclMon). It consists of a simple camera (a) centered above the
eclosion plate, on which the pupa are glued (b). Illumination comes
from the light plates (c, c’) below. One light plate gives white
light for entrainment (c), the other gives constant red light
(λ=635 nm/ λ=850; c’) as illumination for the camera.
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Chapter I
45
2.2 FiJi macro
For analysis of the recorded images and eclosion detection, a
macro for ImageJ/FiJi was
developed by Martin Fraunholz, Microbiology, University of
Würzburg. It is based on the fact
that after eclosion, the fly leaves behind an empty puparium,
which is more light-permeable
(“brighter”) than a puparium with a pharate fly inside (and thus
“darker”). An eclosion event
can therefore be identified by a change of intensity values of
the illumination coming from
below.
The recorded images in the png format were opened as image stack
in FiJi, converted to 8-bit
grey scale and the contrast was enhanced (saturated=10). To
minimize effects of different
light conditions, the background was subtracted from each image
using the rolling ball
algorithm (diameter: 4, light background). Each pupae was
identified by setting of the intensity
threshold and subsequent particle analysis (size range 10 pixels
to infinity) after conversion
into binary images. Square regions of interest (ROI) with a
side-length of 15 pixels were
defined around the center of each particle and added to the ROI
man